REVIEW: Intracellular Cargo Transport by Kinesin-3 Motors

N. Siddiqui and A. Straube*

Received February 1, 2017; Revision received April 11, 2017
Intracellular transport along microtubules enables cellular cargoes to
efficiently reach the extremities of large, eukaryotic cells. While it
would take more than 200 years for a small vesicle to diffuse from the
cell body to the growing tip of a one-meter long axon, transport by a
kinesin allows delivery in one week. It is clear from this example that
the evolution of intracellular transport was tightly linked to the
development of complex and macroscopic life forms. The human genome
encodes 45 kinesins, 8 of those belonging to the family of kinesin-3
organelle transporters that are known to transport a variety of cargoes
towards the plus end of microtubules. However, their mode of action,
their tertiary structure, and regulation are controversial. In this
review, we summarize the latest developments in our understanding of
these fascinating molecular motors.
KEY WORDS: molecular motors, microtubule-based transport,
kinesin, autoinhibition, intracellular transport, Unc104/KIF1, cargo
trafficking

DOI: 10.1134/S0006297917070057

Kinesins are molecular motors that step along microtubule tracks,
thereby converting the chemical energy of one ATP per step into
mechanical work. While moving along the microtubule, kinesins haul
intracellular cargo such as chromosomes or mitochondria to achieve
their correct positioning and transport secretory vesicles from the
cell center to the cell cortex. Common to all kinesins is a structure
that consists of a motor domain, a neck, and a tail. The motor domain
combines both microtubule binding and ATPase activity. The ATP
hydrolysis cycle is coupled to conformational changes within the motor
and neck domains that result in forward movement of the tail-attached
cargo. ATP turnover drives a sequence of conformational changes that
cyclically change the microtubule binding affinity of the motor domains
[1]. Kinesin motors exist in all eukaryotes and
have been divided into 15 families based on the position and sequence
homology of their motor domain [2, 3]. Amongst the 45 human kinesins, the largest family
is the kinesin-3 family, a class of plus-end-directed transporters that
have been implicated in the long-distance transport of vesicles and
organelles in a variety of eukaryotic cells.

The founding member of the kinesin-3 family is Unc-104 from the nematode
worm Caenorhabditis elegans. Mutations in Unc-104 cause impaired
transport of synaptic vesicles to the axon terminal and uncoordinated
and slow movement of the nematode [4, 5]. Kinesin-3 family members have since been
identified as fast organelle transporters in the amoeba
Dictyostelium discoideum [6], as endosome
transporters in fungi [7-9],
and as transporters of vesicles, viral particles, and mitochondria in
mammalian cells [10-17].
Kinesin-3s are thought to have been present in the last common
eukaryotic ancestor, suggesting that cytoplasmic vesicle transport is
evolutionarily ancient, even though today’s land plants lack
kinesin-3s [3]. The kinesin-3 family comprises six
subfamilies: the KIF1, KIF13, KIF14, KIF16, and KIF28 motors [18] plus a fungal-specific group of short
kinesin-3-like proteins [19] (Fig. 1). While vertebrates usually have nine kinesin-3
genes with one to three of these representing each of the five major
subfamilies [18], filamentous fungi usually have
one KIF1 representative (Kin3 in Ustilago maydis, UncA in
Aspergillus, and NKin2 in Neurospora crassa) plus one
short kinesin-3-like protein (UncB in Aspergillus and NKin3 in
Neurospora crassa) [7-9,
19]. Please see Fig. 1 for the
phylogenetic relationship of kinesin-3 family members from vertebrates
(Homo sapiens), insects (Drosophila melanogaster), worms
(Caenorhabditis elegans), and several fungi, incorporating all
motors mentioned in this review.

The number and variety of kinesin-3 motors in higher eukaryotes likely
reflects the requirement for many different cargoes to be transported
into different regions of the cell; thus, the different kinesin-3s are
equipped with different specificities for both the cargo and the
microtubule track and are activated by different mechanisms, as we will
detail in the following sections.

CELLULAR FUNCTION AND HUMAN DISEASE

Kinesin-3-mediated transport is required for neuronal morphogenesis and
function; mutations in any of the KIF1 motors KIF1A, KIF1B, or KIF1C
cause neurological disorders, spastic paraplegia, or multiple sclerosis
both in human patients and mouse models [13, 20-23]. DmUnc104 mutants
also show defects in neuronal development, in particular in the
morphogenesis of synaptic terminals and dendrites [24]. In fungi, transport of endosomes by kinesin-3
motors is required for optimal hyphal growth [9, 25]. Caenorhabditis elegans worms require
axonal transport by Unc-104 for the coordination of their movement [5]. In addition, kinesin-3 motors have been shown to
regulate signaling processes and the orderly progression of cell
division. For example, KIF16A tethers the pericentriolar material (PCM)
to the daughter centriole during mitosis, thereby preventing PCM
fragmentation and enabling the formation of a bipolar mitotic spindle
[26]; and KIF13A translocates a component of the
cell abscission machinery to the spindle midzone, thereby controlling
cytokinesis [27]. Likewise, deletion of the sole
kinesin-3 in U. maydis results in a cell separation defect [7]. Important cargoes of kinesin-3 proteins are
summarized in the table and range from mitochondria and viruses to
vesicles containing a variety of receptors, pre-synaptic signaling
proteins, microtubule regulators, and phospholipids [11, 12, 14,
15, 28-31]. It is becoming clear that the main function of
kinesin-3 motors across species is the long-distance transport of
membranous cargo. Kinesin-3 motors are exceptional in their high
processivity, i.e. the distance they walk before falling off the
microtubule track. This makes them particularly suited for long-haul
tasks, and in the next section we will discuss the structure of
kinesin-3 molecules and point out the features that underlie their
properties.

Kinesin-3 cargoes. List of selected cargoes identified to be transported
by kinesin-3 family members

STRUCTURE OF KINESIN-3 MOTORS

All kinesin motors that walk towards the plus end of microtubules have
their motor domain at the N-terminus of the molecule. This is also true
for kinesin-3 family motors (Fig. 2a). What
sets kinesin-3 motors apart from other kinesins is the organization of
the neck region, which contains a β-sheet as well as a helix [18], and the presence of a forkhead-associated (FHA)
domain [45] in the tail. In addition to the FHA
domain, the tail region contains several short coiled-coils and diverse
protein and lipid interaction domains that mediate binding to cargo and
regulators (Fig. 2, a and b). In this section,
we will discuss kinesin-3 specific features of each region, those that
are common to most kinesin-3 motors and those that give a motor unique
properties.

Fig. 2. Structure of kinesin-3 motors. a) Primary structure of
human kinesin-3 members with characteristic N-terminal motor domain,
FHA domain, and tail with several short coiled-coil (CC) regions in
addition to a variety of protein or lipid interaction motifs. b)
Schematic representation of a dimeric kinesin-3 motor and its
interaction with the microtubule surface as well as a cargo vesicle. c)
Structural model of kinesin motor domains binding to the microtubule
(one αβ-tubulin heterodimer shown, in gray). The flexible
C-terminal tubulin tails (E-hooks) are indicated in green. Key regions
of the kinesin motor domain (blue) that contribute to interaction with
microtubules are highlighted in red for both KIF5A, a kinesin-1, and
KIF1A, a kinesin-3. Key residues that were shown to contribute to
10-fold higher processivity of kinesin-3 are shown in magenta [53, 54]. PDB accession numbers:
4UXP and 4UXY.

The motor domain binds to the microtubule, and the energy from ATP
hydrolysis is used to produce directional movement [46, 47]. A characteristic
feature of the kinesin-3 family is the presence of a stretch of
positively charged lysine residues designated as the K-loop in loop 12
of the motor domain. This loop is ideally positioned so that it can
contact the negatively charged glutamate-rich (E-hook) C-terminal tail
of β-tubulin (Fig. 2c). The K-loop was
proposed to enable processive motion by monomeric KIF1A by mediating
diffusive interaction to microtubules throughout the ATPase cycle [48, 49]. However, while the
K-loop in KIF1, KIF13, and KIF16 has been shown to increase microtubule
affinity [50-52], an increase
in processivity could not be attributed to the K-loop when these motors
are working as dimers [50]. Instead, the K-loop
increases the microtubule-binding rate and enables kinesin-3 motors to
effectively work in teams [50, 51]. Recent comparative high-resolution cryoelectron
microscopy structures of kinesin-1 (KIF5A) and kinesin-3 (KIF1A) motor
domains bound to microtubules in different nucleotide states paired
with molecular dynamics simulations ascertained which family-specific
residue changes result in the 200-fold increased affinity of kinesin-3
motors to microtubules relative to kinesin-1 [53,
54]. These residues reside in loops L2, L7, L8,
L11, L12, and α-helices α4 and α6 (Fig. 2c). Thus, the contribution of multiple sites
increases kinesin-3s’ interaction surface with microtubules and
results in a large effect on affinity. This increased affinity then
increases the processivity of dimeric kinesin-3 motors [54]. Key residues that result in a 10-fold increased
processivity of kinesin-3 versus kinesin-1 are Arg167 in loop 8, Lys266
in loop 11, and Arg346 in α-helix 6 of KIF1A (Fig. 2c) [53].

Coiled coils are important structural features that mediate motor
dimerization [55]. Kinesin-3 motors tend not to
contain the extended coiled coils that are typical for the tails of
other kinesins, but instead contain several smaller predicted
coiled-coil regions scattered along the tail (Fig. 2a). It is presently unclear whether all of these
contribute to dimer formation. So far, the only direct test of this was
performed with the fourth coiled-coil domain of KIF1C, which is
sufficient to drive dimerization in a yeast-two-hybrid assay [56]. In KIF1A, KIF13A, and KIF13B, the coiled-coil
domains seem to interfere with dimerization. It has been shown that
instead, the neck coil alone efficiently dimerizes these motors [57, 58].

FHA domains are small protein modules that recognize phosphothreonine
epitopes on proteins and mediate protein–protein interactions [59, 60]. FHA domains have been
found in more than 200 different proteins with diverse cellular
functions such as transcription, DNA repair, and protein degradation
[61]. Besides fulfilling a structural role in
kinesin-3 proteins, the FHA domain also confers specific cargo
interactions. For example, the FHA domain of KIF13B medicates binding
to its cargo transient receptor potential vanilloid 1 (TRPV1).
Interestingly, this interaction depends on phosphorylation of KIF13B at
T506 in the FHA domain by cyclin-dependent kinase 5 (Cdk-5) [40]. A point mutation that is likely to alter the
folding of the FHA domain of KIF1C causes a change in the
susceptibility of mice to anthrax lethal toxin, further demonstrating
the functional importance of the domain [61, 62].

Several kinesin-3 tails contain domains that allow direct interaction
with membranes, e.g. KIF16A contains a START/lipid sterol-binding
domain at the C-terminus [26]. KIF1A and KIF1B
have a pleckstrin homology (PH) domain that is important for binding
cargo vesicles [63], probably through specific
interaction with phosphatidylinositol 4,5-bisphosphate
(PtdIns(4,5)-P2) [64]. KIF16B possesses
a phosphoinositide-binding structural domain (PX), which binds to
PtdIns(3,4,5)P3 and is involved in the trafficking of early
endosomes [65, 66].

Other kinesin-3 tails contain protein interaction domains, such as a
CAP-Gly domain at the C-terminus of KIF13B. CAP-Gly domains bind to
sequence motifs at the C-terminus of tubulin and EBs, zinc-finger
motifs, and proline rich sequences [67]. KIF1C
possesses a proline-rich region at the C-terminus. Proline-rich regions
play a structural role and also act as binding sites for protein
interaction [68]. In the case of KIF1C, this
domain mediates several protein interactions, including the cargo
adapter protein BICDR1, 14-3-3 proteins, and Rab6 [56, 69, 70].

Surprisingly, a monomeric motor construct of KIF1A has been observed to
undergo processive plus-end directed movement along microtubules [48]. This is thought to be possible due to the
presence of the K-loop and a stable microtubule interaction surface
that persists throughout the ATPase cycle (Fig. 2c) [49, 54]. However, monomeric KIF1A only moves very slowly
(0.15 µm/s) and weakly (~0.15 pN) along microtubules, while
multiple KIF1A motors transport cargo at 1.5 µm/s [14, 71]. Teams of 10 monomeric
KIF1A motors have been proposed to become approximately 100-fold
stronger than a single monomeric motor [72];
however, experimental data on the force generation of kinesin-3 teams
are lacking. There is evidence suggesting that kinesin-3 motors exist
as inactive monomers in cells until activated by dimerization [58, 73-75].
Other studies suggest that KIF1A motors are dimeric in vivo, but
in an autoinhibited state until activated by cargo binding [6, 57]. Thus, the extent to which
individual kinesin-3 family members exist as monomers or dimers in
cells remains to be elucidated. However, it is clear that a single
monomeric motor cannot achieve the high processivity of kinesin-3
mediated cargo transport observed in cells. Thus, these would need
either to work in teams formed by recruitment of several monomeric
motors to the same cargo, or to form dimers.

MECHANISM OF AUTOINHIBITION

Early work and biochemical characterization of conventional kinesin
revealed that the molecule exists in two conformations: a folded
inactive conformation and an extended active one [76, 77]. A small peptide region
in the tail of kinesin-1 binds to the motor domain to inhibit it [78-80]. While kinesin-3 motors
do not contain such an extensive coiled-coil region with a hinge that
allows neat folding and unfolding of the tail, inactive kinesin-3
motors have been shown to adopt a compact conformation with a crumpled
tail [81], which probably extends when activated
and/or under load. That the pool of motors exists in an autoinhibited
state in cells is important because in the absence of cargo, motor
activity needs to be tightly regulated to avoid microtubule crowding
and futile ATP consumption.

Currently, there are two models of autoinhibition that apply to
kinesin-3 motors. In the monomer–dimer switch model,
intramolecular interactions involving neck and tail regions hold some
kinesin-3 motors in a monomeric, inactive state. Upon activation, these
motors dimerize with their neck coil and tail regions undergoing
intermolecular interactions. In the alternative tail block model, the
motors are stable dimers, but regions of the tail interact with the
motor or neck domains and interfere with motor activity until cargo
binding occupies the tail region and releases the motor. Evidence
exists for both models, and the picture emerging is that different
kinesin-3 motors might use either or a combination of both of these
methods of autoinhibition.

Most KIF1 and KIF13 motors are thought to undergo a monomer–dimer
switch. Consistently with an autoinhibited state, the full length
CeUnc-104 and MmKIF1A are inactive in motility assays [14, 57]. As a monomeric motor
domain construct of KIF1A could produce some directional motion by
itself and work as a processive motor when dimerized artificially [48, 75], regions of the neck or
tail interfere with motor activity. Indeed, in Unc-104, the two neck
helices can form an intramolecular coiled-coil, thereby inhibiting the
ATPase and microtubule binding cycle of the motor and holding the motor
in a monomeric state [73]. The neck helices can
also form an intermolecular coiled-coil, thereby enabling the switch
from monomer to dimer, which is required to obtain a processive Unc-104
motor [73]. In MmKIF1A, a similar switch
through intra- and intermolecular coiled-coil formation is proposed to
occur between the neck coil region and the first coiled-coil domain
(CC1). Surprisingly, the truncation of the entire tail results in
processive dimeric motors of KIF1A, KIF13A, and KIF13B, while all
longer constructs containing CC1 result in monomers that only show
diffusive movement [57, 58].
If autoinhibition is prevented by deletion of the flexible hinge
between the neck helices in C. elegans Unc-104, the motility of
the motor in vitro is unperturbed, but transgenic worms show
severe defects in the coordination of their movement [73]. Likewise, mutations in the CC1 segment of KIF1A
result in activation of the motor [82, 83]. In the KIF13 subfamily, a proline residue at the
junction of neck coil and CC1 provides the flexibility to enable CC1 to
fold back and interact with the neck coil. Deletion of this proline
residue results in dimerization via the neck coil domains and active,
processive motors [58, 84].
Control of the autoinhibited state of the KIF1A motor might also
involve the FHA domain and the following coiled coil CC2. A tandem
construct of CC1 and FHA domains forms a very stable dimer.
Furthermore, the dimerization of CC1–FHA sequesters the CC1
region and makes it unavailable for the autoinhibition of the neck coil
region [82]. Also, CC2 can fold back to interact
with the FHA domain, which disrupts the motor activity [57]. Disruption of the CC1–FHA dimer severely
impairs synaptic vesicle transport and locomotion in C. elegans
worms, suggesting that robust dimerization is crucially important for
KIF1A function in vivo [83].

Evidence for a tail-block mechanism exists for KIF13B and KIF16B. In
KIF16B, microtubule binding is inhibited by the interaction of the
second and third coiled coil with the motor domain in an ATP-dependent
manner. This tail-mediated inhibition is important for the correct
localization of early endosomes to somatodendritic regions in neurons
and the recycling of AMPA
(α-amino-3-hydroxy-5-methyl-4-isoxazolpropionate) and NGF (nerve
growth factor) receptors [85]. An interaction of a
tail domain with the motor domain also contributes to the
autoinhibition of KIF13B [38, 86]. Upon phosphorylation of KIF13B close to its
C-terminus by Par1b/MARK2 (microtubule affinity-regulating kinase),
14-3-3β binds and promotes the intramolecular interaction of
KIF13B motor and tail domains. This in turn negatively regulates KIF13B
microtubule binding, resulting in the dispersal of the motor in the
cytoplasm and a reduction in cell protrusion and axon formation [86]. In addition, KIF1C, which is known to exist as a
stable dimer, interacts with 14-3-3 proteins in a
phosphorylation-dependent manner [56]. However,
whether this mediates an autoinhibitory tail–motor interaction
similarly to KIF13B remains to be elucidated.

Taken together, these data suggest specific autoinhibition mechanisms
for each kinesin-3 family member. These might require different
interaction partners to achieve release from autoinhibition and
activate the motors for transport of specific cargoes.

ACTIVATION BY CARGO INTERACTION

Many kinesins are activated upon cargo binding. Full-length KIF13B, also
known as guanylate kinase-associated kinesin (GAKIN), exists in an
autoinhibited state in solution. It is activated by the direct binding
of its cargo, human disc large (hDlg) tumor suppressor [38]. In contrast to KIF1A, full-length KIF13B is
active in a gliding assay. This could be because the binding of the
C-terminus to the glass surface might mimic the cargo-bound state, thus
relieving autoinhibition [38]. In contrast, KIF16B
is a monomer in the cytoplasm and dimerizes at the cargo surface. The
localized dimerization of KIF16B on early endosomes has been directly
observed using Förster resonance energy transfer (FRET) in live
cells [58]. Thus, these examples support the idea
that due to the diverse cargo binding tail, the different kinesin-3
family members use diverse means of autoinhibition and cargo-dependent
release of inhibition, involving changes in the dimerization status for
some members and competitive binding of a peptide region that weakly
interacts with the motor domain for others. The mechanisms of
cargo-mediated activation thus require elucidation for each family
member.

While some motors bind their cargo directly, often cargo adapter
proteins mediate both the motor activation and cargo recruitment. For
C. elegans kinesin-3 motor Unc-104, a number of adapter proteins
are known that are involved in cargo loading; a bimolecular
fluorescence complementation assay (BiFC) was employed to show that
binding of different adapters Unc-16 (JIP3), DNC-1 (DCTN-1/Glued), and
SYD-2 (Liprin-α) to Unc-104 results in translocation to different
subcellular compartments in neuronal cells. This suggests that adapter
proteins can recruit the motor to their cargo and steer their transport
[16, 87]. Further, binding of
LIN-2 (CASK) and SYD-2 was shown to positively regulate the Unc-104
motor by increasing its velocity, and binding of LIN-2 also increased
run lengths. The cargo transport of synaptobrevin-1 (SNB-1) was
markedly reduced in the neurons of LIN-2 knockout worms, implying that
LIN-2 is an activator of Unc-104 motor [88]. In
Ustilago maydis, the cargo adapter Hook protein (Hok1) mediates
the recruitment of Kin3 and dynein to early endosomes and regulates
bidirectional motility. Hok1 releases Kin3, and this allows for dynein
to bind and drive the subsequent change in directionality [41]. Like Kin3, KIF1C binds to another dynein adapter
protein, Bicaudal-D-related protein 1 (BICDR-1) [69]. BICDR-1 also binds Rab6A vesicles, thus linking
both motors to secretory vesicles and controlling the bidirectional
vesicle transport in developing neurons [69].
Centaurin α1 (CENTA1) acts as a cargo adapter for KIF13B and
recruits the motor to
PtdIns(3,4)P2/PtdIns(3,4,5)P3-containing vesicles
[30, 89]. CENTA1 contains two
PH domains that bind the headgroups of phosphoinositides, and PH1 also
directly binds the FHA domain of KIF13B in a
phosphorylation-independent manner [89]. As KIF13B
FHA simultaneously interacts with the ArfGAP domain of a second CENTA1
molecule, CENTA1–KIF13B form a heterotetrameric transport complex
for PtdIns(3,4,5)P3-rich vesicles [30,
89].

REGULATION BY Rab GTPases

The members of the Rab family of GTPases are known to control the
localization of vesicles/organelles in a nucleotide-dependent manner.
Rab proteins act at all stages including vesicle formation, motility,
and tethering of vesicles to the designated compartment [90]. Rab GTPases exist in either GTP or GDP bound
states, and are activated by GEFs (GTP/GDP exchange factors) and
switched off by GAPs (GTPase activating factors) [91]. Once activated, the Rab proteins bind to
vesicles that are translocated to the destination compartment, where
they dock and fuse. The Rab proteins are then recycled back via a
cytosolic intermediate [92]. KIF1A and KIF1Bβ
both transport Rab3-coated vesicles in the axon. Rab3 is a synaptic
vesicle protein that controls the exocytosis of synaptic vesicles [93-95]. It has been found that
DENN/MADD (differentially expressed in normal and neoplastic cells/MAP
kinase activating death domain), a GEF for Rab3, binds to Rab3 and the
tail domain of KIF1A and KIF1Bβ and is thought to mediate the
transport to the axon terminal while maintaining Rab3 in the GTP-bound
form [13].

Rab6 binds to KIF1C at two sites, to the motor domain and near the
C-terminus. Rab6 binding to the motor domain disrupts the motor’s
ability to bind microtubules [70], while the
binding to the C-terminus might activate cargo loading and relief from
autoinhibition. Secretory Rab6 vesicles are transported
bidirectionally, and it is thought that the dual ability of Rab6 to
activate and inhibit KIF1C might regulate the directional switch. KIF1C
also transports Rab11-positive vesicles for the recycling of integrins
[15]. Whether Rab11 is directly involved in
controlling the activity of KIF1C is yet unclear.

The microtubule tracks on which kinesin motors walk are not uniform.
Depending on the cell type or its differentiation status, cells express
different tubulin isoforms, accumulate microtubules with different
posttranslational modifications, and also express different
microtubule-associated proteins (MAPs) that decorate the microtubules.
Kinesins are known to be sensitive to both changes to tubulin and MAP
composition.

Tubulin undergoes a diverse range of chemical modifications known as
posttranslational modifications after polymerization into microtubules.
These modifications mainly occur on the C-terminal tails of both
α- and β-tubulin and include the removal of terminal amino
acids, such as detyrosination, and the addition of polyglutamate and
polyglycine side chains [98-100]. Considering that the kinesin-3-specific K-loop
is thought to interact with the C-terminal tail of β-tubulin
(Fig. 2b), it is expected that changes in this
region would impact kinesin-3 binding. Further modification at other
sites of tubulin have been described, such as the acetylation of K40 in
α-tubulin and phosphorylation of tubulin at various sites [101]. These modifications may change the stability
of microtubules and act as signposts for motor transport by selectively
increasing or decreasing the affinity of certain motors to the
microtubule [102]. In line with this idea,
knockdown of polyglutamylase PGs1 in ROSA22 mice decreases the
localization of KIF1A to neurites [103]. Further,
the ciliary localization of kinesin-3 KLP-6 in C. elegans is
positively regulated by tubulin deglutamylase CCPP-1 [104]. However, in COS cells, the truncated,
constitutively active KIF1A(1-393) was a non-selective motor [105]. The fungal kinesin-3 UncA from A.
nidulans has been reported to selectively walk on detyrosinated
microtubules, and the tail is necessary and sufficient for this
recognition [9, 106]. Also,
the N. crassa kinesin-3 NKin2 preferentially binds to a subset
of microtubules [8]. However, this feature is not
conserved in all fungi, as Kin3 from U. maydis uses all
microtubules equally [107]. Like the finding in
COS cells, the negative result could be due to the lack of modified
microtubules in these cells rather than a different property of the
motor, and this would require further investigation to elucidate. The
subcellular localization of KIF1C is regulated by acetylation in
primary human macrophages in a way that suggests that tubulin
acetylation is a negative signal for KIF1C transport [108]. Likewise, KIF1Bβ and KIF1A have been
reported to drive lysosomal transport preferentially along tyrosinated
(i.e. non-modified) microtubules [109]. These
data suggest that most kinesin-3 motors are sensitive to tubulin
posttranslational modifications, but with different preferences.

MAPs regulate the assembly and disassembly kinetics of microtubules as
well as the interactions of motors with microtubules [110, 111]. Lattice-decorating
MAPs such as the neuronal protein tau regulate the attachment rate and
can act as roadblocks that affect motors differently, depending on
their ability to take side or backwards steps to circumvent the
roadblock [112-114]. MAP4,
which is a tau-related protein in non-neuronal cells, negatively
regulates force generation and transport by dynein, but it positively
regulates kinesin-based movement [115, 116]. Thus, MAPs can regulate microtubule-based
transport directionality and access of motors to microtubules.

For kinesin-3, MAPs known to regulate the motor include
doublecortin-like kinase-1 (DCLK-1), which regulates KIF1 transport of
dense core vesicles (DCVs) along dendrites in neurons. DCLK-1
specifically binds to microtubules in dendrites, which acts as a
positive signal to promote dendritic transport of KIF1 cargoes. In the
absence of DCLK-1, KIF1 motors predominantly transport DCVs into the
axon [117]. In C. elegans, the retrograde
motility of Unc-104 was affected in tau/PTL-1 (protein with tau-like
repeats) knockout worms. Unc-104 usually moves bidirectionally, but in
the absence of PTL-1 the motor travels preferentially in anterograde
direction [118]. It is thought that kinesin-3
motors cooperate with dynein for bidirectional motility, so whether
PTL-1 affects Unc-104 directly or negatively regulates dynein to cause
the observed phenotype remains to be elucidated. The microtubule
plus-end tracking protein CLASP is required to stimulate the
trafficking of KIF1C [119]. KIF1C has also been
described to move with growing microtubule plus ends in cells [120]. This could be either due to the preference for
unmodified (i.e. freshly assembled) microtubules [108], or due to its fast transport speed and thus
ability to catch up with the growing microtubule end [51], or due to its interaction with CLASP [119].

COOPERATION OF MOTORS

Kinesin-3s have been implicated in the bidirectional transport of cargo.
This means that when a specific kinesin-3 is inhibited or depleted, the
transport of its cargo both towards the plus and the minus end of the
microtubule is impaired [15, 118, 121]. It has been
suggested that kinesin-3 cooperates with dynein in the bidirectional
transport of cargoes, but the mechanism underlying the mutual
activation of these opposite-polarity motors remains to be elucidated
[122]. It has been suggested that cooperation
depends on the opposing force generated, resulting in a mechanical
activation [121]. Other proposed models include a
steric inhibition mechanism whereby the direct interaction of the
opposing motor or accessory protein relieves autoinhibition, and a
microtubule tethering mechanism whereby the opposing motor is in a
weakly bound state and acts as a processivity factor [122]. This is different to the idea of tug-of-war
that has been proposed and reconstituted for kinesin-1 and
dynein-mediated transport, where the motors pull against each other and
the strongest team wins, i.e. the number of motors of each type loaded
to a cargo molecule and the force that each motor can produce determine
the net movement of the cargo [123, 124]. Potential linkers to facilitate cooperation of
dynein and kinesin-3 include Hook and Bicaudal, cargo adapter proteins
that have been identified to interact with both dynein and kinesin-3
tail domains [41, 69, 125, 126]. Interestingly, the
presence of BICD2 increases the force generation and processivity of
dynein/dynactin [127, 128],
demonstrating that these cargo adapter proteins regulate motor activity
and could act as switches to control transport directionality within a
complex containing two opposing motors. Other control mechanisms could
come from accessory proteins such as kinesin-binding protein (KBP),
which has been shown to stimulate KIF1B, but inhibit KIF1A-mediated
bidirectional transport [28, 129]. If the activity of such regulatory proteins
were spatially controlled, this would enable directional switching of
transport complexes in the presence of opposing motors.

Kinesin-3 molecules are important cargo transporters in neuronal cells
that show a number of remarkable features. Their high affinity to the
microtubule surface in all nucleotide states makes them highly
processive motors, ideally suited to drive long-distance transport in
neuronal cells. Their processivity is so high that even monomeric motor
domain constructs show some directional motion. The activity of
kinesin-3 motors is tightly regulated, in some motors via a switch from
monomer to dimer, in others via autoinhibitory interactions of motor
and tail domains. These interactions are relieved by cargo recruitment
or regulated by kinases. Kinesin-3 motors are sensitive to the changes
in the microtubule track and follow signposting modifications such as
posttranslational modifications of tubulin or MAP decoration. Finally,
kinesin-3 motors cooperate with dynein to bring about bidirectional
transport of cargo. Many of these fascinating features remain to be
understood mechanistically. Future work will illuminate this problem
and enable us to appreciate kinesin-3 motor physiology, including the
causation of the disease states arising from mutated kinesin-3
motors.

Acknowledgments

We thank Kristen Verhey (University of Michigan) and Carolyn Moores
(Birkbeck) for useful discussions about kinesin-3 motors, and Andrew
McAinsh and Rob Cross for critical reading of the manuscript.

A. S. is a Prize Fellow of the Lister Institute of Preventive Medicine
and a Wellcome Trust Investigator (200870/Z/16/Z). N. S. is funded by a
Chancellor’s International PhD Scholarship of the University of
Warwick.